PAS kinase

ABSTRACT

The invention provides methods and compositions relating to a novel kinase designated PAS Kinase (PASK). The compositions include isolated polypeptides comprising a native PASK protein or a PASK N-terminal domain and polypeptides consisting of a PASK PAS-A or PAS-B domain, as well as isolated polynucleotides encoding such polypeptides, and expression vectors and cells comprising such polynucleotides. The methods include binding assays comprising the steps of incubating a mixture comprising a subject polypeptide with a ligand under conditions wherein the polypeptide binds the ligand; and detecting binding of the polypeptide to the ligand.

CROSS-REFERENCE TO RELATED APPLICATION

This application is a divisional application of and claims priority under 35 U.S.C.§ 120 to 09/770,170, filed Jan. 26, 2001, having the same title and inventors, now U.S. Pat. No. 6,319,679, which is incorporated herein by reference.

INTRODUCTION

1. Field of the Invention

The field of this invention is protein binding assays.

2. Background of the Invention

We have identified a novel enzyme designated PAS-kinase (PASK). The human version of PASK is encoded by a single gene located on chromosome 2, band q37. Analysis of genomic and cDNA clones of PASK has shown the gene to be composed of 18 coding exons covering roughly 37 kilobases of human chromosome 2. The enzyme is 1,323 amino acid residues in length and highly related in primary sequence to polypeptides specified in the genomes of flies and yeast. RT-PCR assays conducted on multiple tissues of the mouse indicate that PASK mRNA is expressed at a similar level in all tissues.

As shown in FIGS. 1A–1C herein, the polypeptide sequences of the human, fly and yeast versions of PASK are most conserved in three regions. Two of the three conserved regions correspond to PAS domains, designated PAS-A and PAS-B, the former being located closer to the amino terminus of the enzyme. The third conserved region of PASK encodes a serine/threonine kinase domain. When compared with all entries available in the database of protein kinases kept by Tony Hunter [1], PASK is most similar in primary amino acid sequence to the catalytic domains of the 5′ AMP activated protein kinase (AMPK), its yeast ortholog SNF1, and the product of the Pim-1 oncogene.

We conducted a two-pronged approach to resolve the biological function of PASK. Biochemical experiments were carried out to study the catalytic activity of the enzyme as well as the modulatory role enacted by the two PAS domains. Complementary genetic experiments were undertaken to study the biological role of PASK in budding yeast. The latter experiments have also directed preliminary studies pertinent to the role of PASK in mammalian cells. In summary, these efforts have established PASK as a serine/threonine kinase that is regulated in cis by its two PAS domains. They have likewise implicated PASK in the regulation of translation and the balance of cell growth (cell size) and mitosis. Biophysical studies of the two PAS domains of PASK have identified PAS-A as a ligand-binding regulatory domain of the PASK enzyme. Together these studies reveal an entirely novel and unanticipated regulatory system that represents a valuable target for the development of synthetic organic compounds capable of regulating the mitotic growth of mammalian cells.

Limited aspects of this disclosure were presented at the 19^(th) International Conference on Magnetic Resonance in Biological Systems at Florence, Italy in August 2000; see, Structural studies of PAS domains: Insight into the link between ligand and protein binding, K H Gardner, C A Amezcua and S M Harper, Aug. 25, 2000, 19^(th) Intrnl Conf Magnetic Resonance in Biological Systems, Florence, Italy; and Solution structure and dynamics of a eukaryotic PAS domain: Evidence for a flexible ligand binding region, C A Amezcua, S M Harper and K H Gardner, Aug. 21, 2000, 19^(th) Intrnl Conf Magnetic Resonance in Biological Systems, Florence, Italy. Genbank accession KIAA0135 has sequence similarity to the disclosed human PASK.

SUMMARY OF THE INVENTION

The invention provides methods and compositions relating to a novel kinase designated PAS Kinase (PASK). The compositions include isolated polypeptides comprising a PASK N-terminal domain, particularly a native PASK protein, and polypeptides consisting of, or consisting essentially of, a PASK PAS-A or PAS-B domain, as well as isolated polynucleotides encoding such polypeptides, and expression vectors and cells comprising such polynucleotides.

The subject methods include binding assays comprising the steps of incubating a mixture comprising a subject polypeptide with a ligand under conditions wherein the polypeptide binds the ligand; and detecting the binding of the polypeptide to the ligand. In particular embodiments, the assay is a kinase assay wherein the ligand is a substrate, the mixture further comprises a nucleoside triphosphate, the binding effects phosphorylation of the substrate, and the detecting step comprises detecting the phosphorylated substrate. In other embodiments, the assay is a NMR-based assay wherein the detecting step comprises detecting an NMR shift in the mixture.

BRIEF DESCRIPTION OF THE FIGURES

FIG. 1A shows an alignment of the PASK PAS-A domains of human (amino acids 131–237 of SEQ ID NO:2), fly (amino acids 72–177 of SEQ ID NO:4) and yeast (amino acids 460–564 of SEQ ID NO:6 and amino acids 288–397 of SEQ ID NO:8).

FIG. 1B shows an alignment of the PASK PAS-B domains of human (amino acids 341–439 of SEQ ID NO:2), fly (amino acids 280–375 of SEQ ID NO:4) and yeast (amino acids 744–843 of SEQ ID NO: 6 and amino acids 522–621 of SEQ ID NO:8).

FIG. 1C shows an alignment of the PASK kinase domains of human (amino acids 1005–1251 of SEQ ID NO:2), fly (amino acids 582–828 of SEQ ID NO:4) and yeast (amino acids 1102–1354 of SEQ ID NO:6 and amino acids 847–1099 of SEQ ID NO:8). The three phosphorylatable activation loop residues are marked with a star.

DESCRIPTION OF PARTICULAR EMBODIMENTS OF THE INVENTION

The nucleotide sequences of cDNAs encoding native PASK polypeptides from human, D. Melanogaster, and yeast (two variants) are shown as SEQ ID NOS:1, 3, 5 and 7, respectively, and the full translates are shown as SEQ ID NOS:2, 4, 6 and 8, respectively. As described in further detail below, these translates comprise an N-terminal functionality, PAS-A and PAS-B domains and a kinase domain.

The subject polypeptides are isolated or pure: an “isolated” polypeptide is unaccompanied by at least some of the material with which it is associated in its natural state, preferably constituting at least about 0.5%, and more preferably at least about 5%, more preferably at least about 50% by weight of the total polypeptide in a given sample and a pure polypeptide constitutes at least about 90%, and preferably at least about 99% by weight of the total polypeptide in a given sample. The polypeptides and polypeptide domains may be synthesized, produced by recombinant technology, or purified from mammalian, preferably human cells. They may be joined, covalently or noncovalently, with a wide variety of conjugates, including labels, tags, etc., particularly other polypeptide sequences. A wide variety of molecular and biochemical methods are available for biochemical synthesis, molecular expression and purification of the subject compositions, see e.g. Molecular Cloning, A Laboratory Manual (Sambrook, et al. Cold Spring Harbor Laboratory), Current Protocols in Molecular Biology (Eds. Ausubel, et al., Greene Publ. Assoc., Wiley-Interscience, N.Y.) or that are otherwise known in the art.

PASK N-terminal domain polypeptides, including the human PASK N-terminal domain (residues 1–89 of SEQ ID NO:2) and functional fragments thereof, elicit PASK specific antibody in a heterologous host (e.g a rodent or rabbit), etc. Accordingly, these polypeptides provide PASK-specific antigens and/or immunogens, especially when coupled to carrier proteins (see, e.g Harlow and Lane (1988) Antibodies, A Laboratory Manual, Cold Spring Harbor Laboratory). For example, polypeptides corresponding to PASK-specific N-terminal domain subsequences are covalently coupled to keyhole limpet antigen (KLH) and the conjugate is emulsified in Freunds complete adjuvant. Laboratory rabbits are immunized according to conventional protocol and bled. The presence of PASK-specific antibodies is assayed by solid phase immunosorbant assays using immobilized PASK polypeptides of SEQ ID NO:2, see, e.g. Table 1.

TABLE 1 Immunogenic N-terminal PASK polypeptides eliciting PASK-specific rabbit polyclonal antibody: PASK polypeptide-KLH conjugates immunized per protocol described above. PASK Polypeptide Sequence Immunogenicity SEQ ID NO:2, residues 1–10 +++ SEQ ID NO:2, residues 12–21 +++ SEQ ID NO:2, residues 20–29 +++ SEQ ID NO:2, residues 32–49 +++ SEQ ID NO:2, residues 52–61 +++ SEQ ID NO:2, residues 65–79 +++ SEQ ID NO:2, residues 77–89 +++

The amino acid sequences of the subject polypeptides are used to back-translate polypeptide-encoding nucleic acids optimized for selected expression systems (Holler et al. (1993) Gene 136, 323–328; Martin et al. (1995) Gene 154, 150–166) or used to generate degenerate oligonucleotide primers and probes for use in the isolation of natural PASK polypeptide-encoding nucleic acid sequences (“GCG” software, Genetics Computer Group, Inc, Madison Wis.). PASK polypeptide-encoding nucleic acids are used in expression vectors and incorporated into recombinant host cells, e.g. for expression and screening, transgenic animals, e.g. for functional studies such as the efficacy of candidate drugs for disease associated with PASK-modulated cell function, etc.

The invention also provides nucleic acid hybridization probes and replication/amplification primers having a PASK N-terminal domain encoding cDNA sequence or fragments thereof sufficient to effect specific hybridization thereto (i.e. specifically hybridize with the corresponding PASK N-terminal domain cDNA). Such primers or probes are at least 12, preferably at least 24, more preferably at least 36 and most preferably at least 96 bases in length. Demonstrating specific hybridization generally requires stringent conditions, for example, hybridizing in a buffer comprising 30% formamide in 5×SSPE (0.18 M NaCl, 0.01 M NaPO₄, pH7.7, 0.001 M EDTA) buffer at a temperature of 42° C. and remaining bound when subject to washing at 42° C. with 0.2×SSPE; preferably hybridizing in a buffer comprising 50% formamide in 5×SSPE buffer at a temperature of 42° C. and remaining bound when subject to washing at 42° C. with 0.2×SSPE buffer at 42° C. PASK nucleic acids can also be distinguished using alignment algorithms, such as BLASTX (Altschul et al. (1990) Basic Local Alignment Search Tool, J Mol Biol 215, 403–410).

TABLE 2 Exemplary PASK nucleic acids which hybridize with a strand of SEQ ID NO:1 under Conditions I and/or II. PASK Nucleic Acids Hybridization SEQ ID NO:1, nucleotides 1–24 + SEQ ID NO:1, nucleotides 25–48 + SEQ ID NO:1, nucleotides 49–72 + SEQ ID NO:1, nucleotides 73–96 + SEQ ID NO:1, nucleotides 97–120 + SEQ ID NO:1, nucleotides 121–144 + SEQ ID NO:1, nucleotides 145–168 + SEQ ID NO:1, nucleotides 169–192 + SEQ ID NO:1, nucleotides 193–216 + SEQ ID NO:1, nucleotides 217–240 + SEQ ID NO:1, nucleotides 242–265 + SEQ ID NO:1, nucleotides 391–711 + SEQ ID NO:1, nucleotides 1021–1317 +

The subject nucleic acids are of synthetic/non-natural sequences and/or are isolated, i.e. unaccompanied by at least some of the material with which it is associated in its natural state, preferably constituting at least about 0.5%, preferably at least about 5% by weight of total nucleic acid present in a given fraction, and usually recombinant, meaning they comprise a non-natural sequence or a natural sequence joined to nucleotide(s) other than that which it is joined to on a natural chromosome. Recombinant nucleic acids comprising native PASK cDNAs, or fragments thereof, contain such sequence or fragment at a terminus, immediately flanked by (i.e. contiguous with) a non-native sequence (a sequence other than that which it is joined to on a natural chromosome), or immediately flanked by a native flanking region of fewer than 2000, preferably fewer than 500, more preferably fewer than 100 nucleotides. While the nucleic acids are usually RNA or DNA, it is often advantageous to use nucleic acids comprising other bases or nucleotide analogs to provide modified stability, etc.

The subject nucleic acids find a wide variety of applications including use as translatable transcripts, hybridization probes, PCR primers, diagnostic nucleic acids, etc.; use in detecting the presence of PASK genes and gene transcripts and in detecting or amplifying nucleic acids encoding additional PASK homologs and structural analogs. In diagnosis, PASK hybridization probes find use in identifying wild-type and mutant PASK alleles in clinical and laboratory samples. Mutant alleles are used to generate allele-specific oligonucleotide (ASO) probes for high-throughput clinical diagnoses. In therapy, therapeutic PASK nucleic acids are used to modulate cellular expression or intracellular concentration or availability of active PASK.

The invention provides efficient methods of identifying agents, compounds or lead compounds for agents active at the level of a PASK modulatable cellular function. Generally, these screening methods involve assaying for compounds which modulate the interaction of a subject PASK polypeptide with a ligand and/or natural binding target. A wide variety of assays for binding agents are provided including labeled in vitro protein-protein binding assays, immunoassays, cell based assays, etc. The methods are amenable to automated, cost-effective high throughput screening of chemical libraries for lead compounds. Identified reagents find use in the pharmaceutical industries for animal and human trials; for example, the reagents may be derivatized and rescreened in in vitro and in vivo assays to optimize activity and minimize toxicity for pharmaceutical development.

In vitro binding assays employ a mixture of components including a PASK polypeptide, which may be part of a fusion product with another peptide or polypeptide, e.g. a tag for detection or anchoring, etc. The assay mixtures comprise a ligand, which term is used generically to encompass specific binding targets including substrates, preferably small molecule ligands as opposed to large molecule ligands such as protein ligands. In a particular embodiment, the binding target is a substrate of PASK kinase activity. The assay mixture also comprises a candidate pharmacological agent. Candidate agents encompass numerous chemical classes, though typically they are organic compounds; preferably small organic compounds and are obtained from a wide variety of sources including libraries of synthetic or natural compounds. A variety of other reagents may also be included in the mixture. These include reagents like ATP or ATP analogs (for kinase assays), salts, buffers, neutral proteins, e.g. albumin, detergents, protease inhibitors, nuclease inhibitors, antimicrobial agents, etc. may be used.

The resultant mixture is incubated under conditions whereby, but for the presence of the candidate pharmacological agent, the PASK polypeptide specifically binds the ligand with a reference binding affinity. The mixture components can be added in any order that provides for the requisite bindings and incubations may be performed at any temperature which facilitates optimal binding. Incubation periods are likewise selected for optimal binding but also minimized to facilitate rapid, high-throughput screening.

After incubation, the agent-biased binding between the PASK polypeptide and one or more ligands is detected by any convenient way. For PASK kinase assays, ‘binding’ is generally detected by a change in the phosphorylation of a substrate. In this embodiment, kinase activity may be quantified by the transfer to the substrate of a labeled phosphate, where the label may provide for direct detection as radioactivity, luminescence, optical or electron density, etc. or indirect detection such as an epitope tag, etc. A variety of methods may be used to detect the label depending on the nature of the label and other assay components, e.g. through optical or electron density, radiative emissions, nonradiative energy transfers, etc. or indirectly detected with antibody conjugates, etc.

A difference in the binding affinity of the PASK polypeptide to the ligand in the absence of the agent as compared with the binding affinity in the presence of the agent indicates that the agent modulates the binding of the polypeptide to the ligand. A difference, as used herein, is statistically significant and preferably represents at least a 50%, more preferably at least a 90% difference.

The following experimental section and examples are offered by way of illustration and not by way of limitation.

EXAMPLES

Biochemical properties of the Pask enzyme. A recombinant baculovirus was engineered to overexpress human PASK enzyme in cultured insect cells. The enzyme was expressed as a fusion with a His₆ tag at its amino terminus such that the recombinant polypeptide could be purified by Ni-affinity chromatography. Routine enzyme preparations were harvested from 0.8 liter of cultured Sf-9 cells. Cells were lysed by dounce homogenization in 50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 0.04% β-mercaptoethanol (“lysis buffer”) and spun at 100,000×G for one hour to remove insoluble debris. Soluble material (˜50 ml) was then applied to a 3 ml bed volume of Ni-affinity resin. Following application and 50× column volume wash, bound material was eluted with storage buffer (50 mM HEPES, pH 7.4, 100 mM NaCl, 1 mM DTT, 1 mM EDTA) supplemented with 250 mM imidazole. Eluted material was sequentially subjected to Mono-Q, anion-exchange chromatography, and eluted with increasing NaCl concentration. Fractions were evaluated by SDS-PAGE to identify PASK enriched fractions, pooled and dialyzed against storage buffer supplemented with 10% glycerol. Typical enzyme preparations yield 0.5 mg of intact PASK enzyme judged to be greater than 95% pure by visual inspection of Coomassie-stained SDS-PAGE gels.

Purified PASK was observed to become phosphorylated when exposed at room temperature to ATP and magnesium. Protein dilution studies revealed that the level of PASK phosphorylation diminished significantly upon dilution, indicating that phosphorylation occurs in trans. As will be shown subsequently, phosphorylation significantly elevates the activity of PASK by enhancing its K_(m) for substrate without altering the catalytic rate constant of the enzyme. Mass spectrometry was employed to identify the sites of auto-phosphorylation suffered by PASK in response to ATP. The major sites of phosphorylation are restricted to two short sequences, one corresponding to the putative activation loop of the catalytic domain and another corresponding to a serine-rich segment of the polypeptide located on the immediate carboxyl terminal side of the catalytic domain. Roughly 2 moles of phosphate were observed to modify a tryptic peptide encompassing the putative activation loop of PASK, and an additional 3–4 moles of phosphate were observed to modify the C-terminal, serine-rich segment of the enzyme.

Kinetic data were used to characterize the activities of PASK enzyme samples derived without modification from baculovirus and following pre-phosphorylation in the presence of ATP. Enzyme activity was monitored using a synthetic substrate bearing the sequence NH₂-AMARAASAAALARRR-CO₂H (AMARA peptide, SEQ ID NO:9). The serine residue in this peptide is known to be phosphorylated by both AMPK and Snf1 [2], both of which are exceedingly similar in primary amino acid sequence to the catalytic domain of PASK. The three arginine residues located at the carboxyl terminus of the AMARA peptide allow avid binding to phosphocellulose paper, thereby facilitating quantitation of ³²P incorporation catalyzed by PASK from [γ-³²P]ATP. Whereas the observed V_(max) for the two enzyme samples was equivalent, K_(m) for substrate differed significantly. The prephosphorylated enzyme was observed to have a 6.5-fold lower K_(m) for substrate than the unmodified enzyme (Table 3). Moreover, lambda phosphatase treatment generated an enzyme which had 7-fold lower activity than the unmodified enzyme, and this treatment was fully reversible by allowing the enzyme to autophosphorylate in the presence of ATP.

Having mapped sites of phosphorylation to two segments of PASK, mutated derivatives of the enzyme were produced as a means of assessing the effects of phosphorylation on enzyme activity. The putative activation loop of PASK can be identified according to extensive studies on AMPK, one of the two closest relatives of PASK. It is known that threonine residue 172, located within the activation loop of AMPK, must be phosphorylated in order to support significant catalytic activity [3]. Mutated derivatives of PASK were generated wherein serine residue 1149, threonine residue 1161 (which aligns exactly with T172 of AMPK), and threonine residue 1165 were individually changed to alanine. All three of these residues map to a tryptic peptide that is autophosphorylated by PASK. As shown in Table 3, the S to A mutation at residue 1149 (S1149A) did not significantly alter either the catalytic rate or K_(m) for substrate. The T to A mutation at residue 1165 (T1165A) completely eliminated all detectable activity. Finally, the T to A mutation at residue 1161 (T1161A) led to the formation of an enzyme that was insufficiently stable to be purified from baculovirus and tested for enzymatic activity. Given the observation that two moles of phosphate were detected by mass spectrometric analysis of the tryptic peptide covering the activation loop of PASK, we tentatively conclude that the fully active enzyme is phosphorylated on threonine residues 1161 and 1165. These data indicate that PASK can, in trans, phosphorylate itself in a manner that mobilizes the activation loop of the enzyme and substantially enhances the avidity of the enzyme for substrate.

TABLE 3 Kinetic constants, K_(m) and V_(max), for wildtype and mutant forms of PASK using peptide substrate. Pre-phosphorylated Untreated Enzyme K_(m) (mM) SE V_(max) SE K_(m) (mM) SE V_(max) SE wildtype 0.2 0.05 0.2 NS 1.3 0.3 0.1 NS S1149A 0.6 0.05 0.1 NS T1161A N/A T1165A Catalytically inactive S1273,77, 0.3 NS 0.4 NS 80A S1287,89A 0.2 NS 0.1 NS S1273- 0.2 NS 0.1 NS 1289A PASK ΔN 0.3 NS 2.8 NS

For this data table, enzyme assays were performed as described herein. PASK concentrations (0.5–2 μg/ml) were determined to have linear rates with respect to protein concentration and time. Kinetic constants were determined using peptide substrate concentrations between 0.06–1 mM. The data were fit to a non-linear Michaelis-Menten equation using the GraphPad Prism 2.0 program. V_(max) values given are μmol/min/mg. NS indicates an SE less than 0.05.

Three additional variants of the enzyme were engineered, expressed and studied in functional assays to assess the potential relevance of phosphorylation in the serine-rich, carboxyl-terminal domain of PASK. One mutant changed serine residues 1273, 1277 and 1280 to alanine (S1273,77,80), a second mutant changed serine residues 1287 and 1289 to alanine (S1287,89A), and a third mutant simultaneously changed all five serines to alanine (S1273-89A). Each of the three mutated enzymes were expressed in baculovirus, purified and tested for both substrate avidity and catalytic rate. All three variants exhibited activities similar to the native enzyme.

A final series of experiments was conducted to investigate the influence of the amino terminal segment of PASK on catalytic activity of the enzyme. The only sequence conservation identifiable among the amino terminal segments of human, fly and yeast enzymes are two PAS domains (FIGS. 1A–1C). A truncated variant lacking both PAS domains (PASK ΔN) was engineered and expressed in baculovirus-infected Sf-9 cells. This variant of the human enzyme removed 948 residues on the amino-terminal side of the catalytic, serine/threonine kinase domain, and was expressed with a His₆ tag at the amino terminus of the truncated polypeptide. Although K_(m) for substrate was unaffected, this truncated variant exhibited a catalytic rate constant (V_(max)) 18-fold higher than the native enzyme (Table 3). In addition to this apparently derepressed basal activity, the truncated enzyme is not subject to regulation by phosphorylation. In contrast to the native enzyme, the lambda phosphatase-treated enzyme and the enzyme pre-incubated with magnesium and ATP showed identical catalytic activity to the unmodified protein. In summary we conclude that the catalytic activity of PASK is negatively regulated by amino terminal segments of the enzyme, particularly corresponding to the two PAS domains of the enzyme. Likewise the K_(m) for substrate of this enzyme can be modified by phosphorylation of two threonine residues corresponding to the activation loop of the kinase domain.

In summary, biochemical studies of PASK have described an enzyme that is fundamentally very similar to the AMPK and Snf1 enzymes. Like PASK, AMPK activity is repressed by the non-catalytic regulatory domain of the enzyme (reviewed in [4]). When the regulatory domain is derepressed by increased AMP concentration, or is removed by engineered deletion, the catalytic rate of the kinase domain is substantially enhanced [5, 6]. The amino acid sequence of the catalytic domain of PASK is more similar to those of the AMPK/Snf1 and Pim1 enzymes than any other proteins in public databases, and like the AMPK/Snf1 and Pim1 enzymes, this catalytic domain is subject to phosphorylation-mediated activation in its activity for substrate. Moreover, the amino terminal domain of PASK, which contains two PAS domains, negatively regulates the catalytic activity of its serine/threonine kinase domain in cis—as does the regulatory domain of AMPK/Snf1. Finally, the PAS domains of PASK can perform a sensing function.

Genetic studies of PASK in budding yeast. The genome of the budding yeast, Saccharomyces cerevesiae, contains genes encoding two highly related forms of the PASK enzyme. One gene, designated PSK1, is located on the left arm of chromosome 1, corresponding to the open reading frame (ORF) YAL017W. The other, designated PSK2, is located on the left arm of chromosome 15, corresponding to ORF YOL045W. The two forms of yeast PASK are more related in primary amino acid sequence to one another than either are to the enzymes encoded by flies or humans. As such, we tentatively conclude that they arose as a duplication subsequent to the evolutionary divergence between yeast and the other metazoan organisms presently under study. The data indicate, however, that the yeast forms of PASK represent bona fide orthologs of the metazoan enzymes. The amino acid sequence of the catalytic domains of the two yeast enzymes are more highly related to those of the metazoan PASK enzymes than any other protein kinase, including all protein kinases encoded by the genome of S. cerevesiae. The same holds for both PAS domains of the two yeast enzymes. The PAS-A domains of both yeast enzymes are more highly related in primary amino acid sequence to the PAS-A domains of metazoan PASK enzymes than any PAS domain available in public databases. Likewise, the PAS-B domains of the yeast enzymes are more highly related to the PAS-B domains of metazoan PASKs than any other PAS domain. Finally, we have obtained unequivocal evidence that human PASK effectively complements the growth defects of yeast strains lacking both PSK1 and PSK2 enzymes (see below).

The individual yeast PSK1 and PSK2 genes were deleted by homologous recombination. In both cases a targeting vector was prepared to replace the entire open reading frame with a selectable marker. Evidence of successful deletion of each gene was confirmed by Southern blotting, yet no obvious phenotypic effect on growth under normal conditions was observed. A doubly-deleted strain (psk1 psk2) was generated by eliminating the PSK2 gene from a haploid, psk1 mutant strain. Again, no obvious effects on vegetative growth were observed. The doubly-deleted strain was then subjected to a wide range of culture conditions in an effort to identify conditions that might reveal differences between wildtype yeast, singly-mutated variants and the doubly-deficient strain. Among 50 prototypical growth conditions tested [7], two were observed to differentially affect the growth of the strain lacking both PSK1 and PSK2 genes. Cultures containing elevated levels of zinc (10 mM), when incubated at 38° C., supported the growth of wildtype yeast to a more substantive degree than the psk1 psk2 double mutant. Under such conditions the PSK2-deleted strain grew at an intermediate level, whereas the PSK1-deficient strain was similar to wildtype. Very similar observations were made when vegetative growth at 38° C. was monitored on minimal-media culture plates with galactose rather than glucose as the carbon source. Under such conditions, the wildtype strain grew well, psk1 cells grew at a slightly reduced rate, psk2 cells grew at a markedly reduced rate, and the psk1 psk2 double mutant grew very poorly. Under both conditions, reintroduction of PSK2 on a plasmid completely restored growth to wildtype levels.

Having established culture conditions capable of distinguishing vegetative growth rate, efforts were made to identify high-copy suppressors of the psk1 psk2 double mutant phenotype. Two independently generated libraries of yeast genomic DNA, carried on a high-copy plasmid bearing the yeast URA3 gene (pRS426) [8], were transformed into the double mutant. Cells were initially selected on minimal media culture plates supplemented with glucose, yet lacking uracil. Three to four days later Ura⁺ colonies were scraped from each dish, pooled and plated onto minimal media culture plates with galactose and incubated at 38° C. Under such conditions, cells of the psk1 psk2 double mutant fail to grow. When transformed with either of the two genomic libraries, however, evidence of high-copy suppression was observed in the form of healthy colonies. Suppressing plasmids were rescued from individual colonies, transformed into E. coli, isolated and sequenced in order to identify putative high-copy suppressors. This effort led to the isolation and identification of roughly 50 high-copy suppressors of the psk1 psk2 double mutant phenotype. Among these, we focused our attention on clones that were retrieved a minimum of two independent times, amounting to fifteen independent genes (Table 4). Each of these genes was retransformed into psk1 psk2 mutants and the suppressing phenotype was confirmed. Two of the suppressing genes are the PSK genes themselves, demonstrating the validity of the screen. Three of the genes encode proteins involved in sugar metabolism; including phosphoglucose mutase 1 (Pgm1p), phosphoglucose mutase 2 (Pgm2p), and Snf1 interacting protein (Sip1p). Pgm1p and Pgm2p are functionally redundant and represent critical enzymes in the conversion of galactose to glucose [9, 10], and Sip1p is a positive regulator of the Snf1p kinase controlling alternate carbon source utilization [11]. A fifth gene repeatedly isolated in the suppressor screen, designated DDP1, encodes diadenosine polyphosphate hydrolase [12, 13].

Among the remaining ten genes that were retrieved at least two independent times in the high-copy suppressor screen, seven—and possibly two others—encode products involved in RNA metabolism or translation. These include: (i) the DED1 and DBP1 genes—both of which encode RNA helicases involved in translation initiation [14–16]; (ii) EDC1—which encodes an enhancer of mRNA decapping [17] (iii) RPR1 and POP4—which respectively encode the RNA and a protein component of RNaseP [18, 19]; (iv) tG(CCC)D which encodes a glycine tRNA [20]; and (v) RDN which encodes the RNA subunits of the ribosome [21]. Additionally, plasmids containing the YDL189W ORF were recovered multiple, independent times in the high-copy suppressor screen. Although no function has been ascribed to this gene, the encoded protein contains an R3H single-stranded nucleic acid binding domain [22], indicating that this gene product may also be involved in RNA metabolism or translation. Finally, the CPA1 gene encoding carbamoyl phosphate synthetase also emerged as a bona fide high-copy suppressor of the psk1 psk2 double mutant phenotype. Although the product of this gene acts at the first catalytic step of arginine biosynthesis [23], Cpa1p translation is tightly regulated via a μ-ORF in its 5′ UTR [24, 25].

TABLE 4 High-copy suppressors of psk1 psk2 mutation. Gene # Description PSK1/PSK2 4 PAS Kinase PGM1/PGM2 3 Galactose Metabolism SIP1 5 Glucose Repression DDP1 4 Diadenosine Polyphosphate Hydrolase DED1 3 RNA Helicase DBP1 2 RNA Helicase EDC1 2 Enhancer OF mRNA Decapping RPR1 2 RNase P RNA Subunit POP4 3 RNase P/RNase MRP Subunit tG(CCC)D 2 tRNA Gly RDN 3 Ribosomal RNA subunits YDL189W 10  R3H Domain Protein CPA1 4 ARG Biosynthesis ADE16 2 Adenine Biosynthesis

For this data table, suppressors were recovered multiple times. The first column gives the gene or ORF name. The second gives the number of independent times this gene was recovered in the suppressor screen. The third column gives a brief description of the gene products of the high-copy suppressor genes.

Our genetic studies of the two yeast genes encoding PASK have helped establish a framework for understanding the biological role of this enzyme. Yeast cells without both enzymes are significantly impeded in vegetative growth when the cells are supplied with sub-optimal sources of sugar. Since seven of the eleven relevant high-copy suppressors—and as many as nine—encode products relating to RNA metabolism or translation, we conclude that PASK is involved in protein synthesis. Moving forward with this observation we prepared a dominant negative form of the human PASK enzyme in which a conserved lysine residue located within the ATP-binding region of the catalytic domain was mutated to arginine (designated K1028R). Baculovirus-expressed K1028R is completely inactive with respect to both autophosphorylation and trans-phosphorylation of the AMARA peptide. An expression vector encoding the K1028R variant of human PASK was transfected into cultured Rat-1 cells. Stable transformants were isolated and evaluated by western blotting using anti-hPASK antibodies. Two cell lines were chosen for further study, one expressing an intermediate level of the K1028R variant of PASK (designated KRint), and another expressing a higher level of the inactive enzyme (designated KRhi). The growth rate of the KRint cell line was slightly slower than that of Rat-1 cells that had been transfected with an empty expression vector. The KRhi cells doubled much more slowly, roughly 25% of the growth rate of control cells.

The KRhi cells were also observed to be substantially larger than control cells when viewed by light microscopy. Following this observation, FACS experiments were conducted to measure both the size and DNA content of log phase KRhi, KRint and control cells. The size distribution of KRhi cells was substantially different from both the KRint and control cells. More dramatic differences were observed in measurements of DNA content. KRhi cells display a bi-modal DNA contcorresponding to 2N and 4N the diploid DNA content of control Rat-1 cells. These observations are consistent with images of DAPI-stained cells. The nuclei of KRhi cells are considerably larger than those of control Rat-1 cells. In summary, these studies indicate that high levels of expression of a dominant-negative form of PASK slows the mitotic growth of Rat-1 cells and leads to a concomitant increase in both cell size and DNA content.

Our identification of a link between PASK—implicated in translational regulation by yeast genetics—and cell size control has a striking parallel to results of Thomas and colleagues [26]. By conditionally eliminating the 40S ribosomal protein S6 in adult mouse liver tissue, the Thomas laboratory has provided compelling evidence that lesions in the translation apparatus differentially affect mitotic growth as compared with cell size. Hepatocytes deficient in the S6 protein are unable to divide mitotically subsequent to hepatectomy. By contrast, when experimental animals are starved, these same S6-deficient hepatocytes are able to increase in cell size in response to re-feeding. Indeed, entire liver mass expands similarly in control and S6-deficient tissue in response to re-feeding. The very surprising and compelling observations of Thomas and colleagues reveal a fundamental difference between growth in cell size and mitotic growth in translationally-compromised hepatocytes.

The observations reported by Thomas and colleagues are consistent with emerging evidence that the genes encoding certain translation initiation factors are able to contribute to the transformed phenotype of cultured mammalian cells. The most compelling evidence that dysregulation of translation can foster transformation has come from studies of the 5′ cap-binding factor eIF4E. Overexpression of this translation initiation factor has been shown to directly induce malignant transformation of NIH-3T3 cells [27]. Additional evidence indicating that enhancement of translation initiation can foster the transformed phenotype has come from studies of eIF4E expression levels in human tumors. As reviewed by Clemens and Bommer[28] and DeBendetti and Harris [29], eIF4E levels are significantly elevated in many solid tumors and tumor cell lines. The most pronounced increases in eIF4E expression have been found in breast cancer tissues, and head and neck squamous cell carcinomas [30–34]. Although evidence favoring the role of enhanced translation in the formation of human tumors is strongest in the case of eIF4E, elevated levels of mRNAs encoding eIF4A and eIF4G have also been detected in melanoma and squamous cell lung carcinoma tumors [35, 36]. Finally, the p48 subunit of eIF3 has been identified as the protein encoded by the int-6 gene, which is a site of frequent integration in the mouse genome by mouse mammary tumor virus [37].

Extensive mechanistic studies have resolved the biochemical functions of both positively- and negatively-acting translation initiation factors. The activity of the eIF4E cap binding factor is negatively regulated by proteins designated eIF4E binding proteins (4E-BPs). As reviewed by Raught and Gingras [38], both eIF4E and 4E-BPs are regulated by phosphorylation. The MAP kinase pathway (growth factor regulated), TOR pathway (growth factor regulated) and p38 pathway (stress regulated) have all been implicated in the phosphorylation of 4E-BPs. When phosphorylated, the 4E-BPs are released from eIF4E, allowing the cap binding protein to interact with eIF4G, form a stable interaction with 5′ cap structures, and enhance the efficiency of translation initiation. Most components of this translational regulatory complex have been shown to be cytoplasmic and many tend to be enriched in the perinuclear region of mammalian cells [39]. Notably, PASK is identically distributed as assayed by immunohistochemical staining of cultured mammalian cells. In sum, these data indicate PASK can act by directly regulating the activity of specific translation initiation factors.

Biophysical studies of the PASK enzyme. In parallel with the functional studies of PASK, we undertook NMR-based studies of the structure, dynamics and ligand binding of the PAS domains of this protein. Our goals were two-fold: to determine if these domains are structurally homologous to other members of the PAS domain family and if so, to investigate whether they are capable of binding small molecule ligands in a similar manner.

Given the requirement for large (10–100 mg) quantities of soluble, isotopically-labeled protein for NMR studies, our first step was to optimize expression of PAS-containing constructs in bacterial systems. To this end, we used secondary structure prediction methods [40] to identify PAS domains from the primary amino acid sequence of PASK and design constructs that avoided having termini within secondary structure elements. These were cloned into a series of “parallel” expression vectors optimized for ease of cloning and expression [41] to rapidly generate a group of plasmids that can use bacteriophage T7-based systems to express the PAS fragments as C-terminal fusions to several partner proteins. As SDS-PAGE analysis identified several soluble fragments of PAS-A, we initially concentrated our attention on those.

We evaluated soluble fragments of PAS-A for their suitability for NMR studies by using a rapid NMR-based screening method [42, 43]. This approach is based on 2D ¹⁵N/¹H HSQC spectra of E. coli cell extracts from uniformly ¹⁵N-labeled cultures that have overexpressed proteins of interest. For proteins overexpressed at the levels we typically observe from T7-promoter driven systems, the high abundance of these proteins in the extract ensures that signals in these spectra are exclusively derived from the protein of interest. From these spectra, we can qualitatively estimate the number of residues in random coil conformations from the amide proton chemical shift dispersion, which is typically poor for unfolded proteins (7.8–8.4 ppm) as compared to folded proteins (6–10 ppm). As no protein purification is required, and these spectra can be acquired in 15–60 minutes on a 500 MHz NMR instrument, this method provides a quick assessment of folding and project feasibility at an early stage. Using this approach for a series of C-terminal deletion constructs of the PAS-A domain, we identified residues 131–237 as a minimal folding unit for this domain.

Having identified a well-folded PAS-A fragment that included residues 131–237, we began structural studies of this domain. Using size exclusion chromatography and ¹⁵N relaxation measurements, we established that this fragment was monomeric and thus could use standard triple resonance approaches to obtain ¹⁵N, ¹³C and ¹H chemical shift assignments. As several reviews of these methods are available [44, 45], we will briefly survey this approach here. All of the NMR experiments were run on 1 mM samples of uniformly ¹⁵N or ¹⁵N/¹³C labeled PAS-A using 500 and 600MHz NMR spectrometers. Backbone chemical shift assignments were based on 3D HNCACB, CBCA(CO)NH, HNCO and HNHA spectra. Sidechain assignments for non-aromatic residues were obtained from 3D H(CCO)NH-TOCSY, (H)C(CO)NH-TOCSY and HCCH-TOCSY methods. Stereospecific assignments of Val and Leu methyl groups were based on constant time ¹³C/¹H spectra acquired on a uniformly 10% ¹³C-labeled sample [46]. Proton chemical shift assignments for the ring systems of the nine aromatic amino acids were based on 2D NOESY, DQF-COSY and TOCSY spectra recorded on an unlabeled PAS-A sample in 99.9% D₂O. Using this combination of methods, we obtained virtually complete chemical shift assignments for PAS-A.

Using these assignments, we determined the solution structure of PAS-A from the following information. Distance restraints were obtained from two isotope-edited 3D NOESY experiments (¹⁵N- and ¹⁵N,¹³C-edited [47]; τ_(m)=150 ms) run on a uniformly ¹⁵ N/¹³C sample in 90% H₂O:10% D₂O and a 2D NOESY spectrum (τ_(m)=120 ms) of an unlabeled sample in 99.9% D₂O. NOE peak intensities were qualitatively interpreted into three distance classes, accounting for chemical shift degeneracy [48]. Restraints on the backbone dihedral angles φ and ψ were derived from analyses of backbone ¹H, ¹⁵N and ¹³C chemical shifts using the program TALOS [49] with minimum bounds of ±30°. Hydrogen bond restraints were derived from qualitative analyses of ²H exchange data, defining protected amides as those with significant protonation after 1 hour in 99.9% D₂O at pH 6.5, 25° C. Structures were generated with a standard simulated annealing protocol in CNS [50] using 515 manually assigned NOE-based distance restraints, 116 dihedral angle restraints and 48 hydrogen bond restraints. These structures were subsequently refined using an ambiguous NOE protocol as implemented in the ARIA enhancement of CNS [51]. Using ARIA-based interpretation of the 2D and 3D NOE spectra, we increased the number of interproton distance restraints from 515 to 2520. Furthermore, 61 ¹⁵N/¹H residual dipolar coupling restraints were added based on measurements made on a sample of PAS-A in a suspension of 10 mg/mL Pf1 filamentous bacteriophage [52].

We generated a group of 20 structures from this information with low total energy and few violations of experimental restraints. The high precision of these structures is indicated in the low backbone r.m.s. deviation of individual structures to the mean: 0.44±0.08 Å for residues located outside the Fα/FG region. This precision is typical of structures with a high average number of restraints per residue, as we have here (˜25 restraints/residue). Most sidechains within the core of the protein are also well defined, with an r.m.s. deviation of 0.95±0.12 Å. Comparing these structures to those of other PAS domains [53–56] show that all have a similar global α/β fold, with a backbone r.m.s. deviation of 1.2–1.5 Å for residues in the β-sheet and N-terminal Cα, Dα and Eα helices.

Comparison of PAS domain structures reveals a key difference between the PAS-A domain of PASK and other PAS domains. In contrast to the long Fα helix observed in other structures, this helix and the subsequent FG loop are significantly disordered in PASK. This is consistent with our observations that signals from these regions were attenuated or missing in several of the spectra used for backbone chemical shift assignment. To investigate this further, we measured ¹⁵N relaxation rates to characterize the backbone dynamics of PAS-A. ¹⁵N T₁, T₂ and heteronuclear ¹⁵N{¹H} NOE values were determined using standard ¹⁵N/¹H HSQC-based experiments [57] and analyzed with a reduced spectral density function mapping approach [58, 59]. These results provide an experimental measurement of flexibility of the protein backbone on several timescales. Most of the protein is well ordered, as judged by the relatively uniform J values in residues in the Aβ–Dα and Gβ–Iβ secondary structure elements. However, the region between Eα and Gβ shows increased J(0.87ω_(H)) and J_(eff)(0) values, consistent with increased flexibility on the nanosecond and millisecond timescales. This contrasts with ¹⁵N relaxation measurements on other PAS domains[55], which do not show any unusual flexibility in any secondary structure elements.

Based on our studies to this point, two lines of evidence pointed to the possibility that the PAS-A domain was capable of binding small ligands. First, the combination of structural and dynamic data clearly demonstrates the flexibility of the Fα and FG regions. Both of these regions are intimately involved in PAS/ligand interactions as shown in the structure of the FixL PAS domain bound to heme [56]. If PAS-A binds ligands in an induced fit manner, it is reasonable that these regions will be flexible in the absence of compound. Second, structure-based alignments show that key positions [60] in PAS-A are occupied by amino acids that may allow small molecules to bind in the core of this domain. These same positions are occupied by large hydrophobes in PAS domains that bind small ligands (PYP [53]) or no ligands (HERG [54]), effectively filling the central core. In contrast, FixL has two glycines in these positions to facilitate binding of the large heme ligand [56]. In PASK these sites are taken by amino acids that may similarly allow access into the core of the domain, due to their small size (Cys) or conformational preferences (Ile).

To directly investigate the ligand-binding capability of the PAS-A domain, we used NMR-based methods to screen a small library of synthetic organic compounds. This library consisted of approximately 150 compounds with formula weights between 200–300Da. Compounds were screened by adding groups of 5 (500 μM each) into samples of uniformly ¹⁵N-labeled protein (250 μM) and monitoring ligand-induced conformational changes in PAS-A by recording ¹⁵N/¹H HSQC spectra on these samples. If significant chemical shift changes were observed in these multiligand samples, we deconvoluted the contribution of each by examining the shifts of each ligand individually in new samples. In this manner, we identified eight compounds that bound PAS-A with K_(d) values (measured by NMR) below 1 mM. Spectra showing the chemical shift changes caused by the tightest binding of these compounds were determined. These data show that titrating this compound into a PAS-A sample leads to chemical shift changes in a limited number of residues that cluster in a region that is highly analogous to the heme-binding site of FixL [56].

We have obtained structure/activity relationship (SAR) information by examining the binding characteristics of structurally related compounds. For example, compound #134 has a bibenzyl architecture, a common feature of many protein-binding ligands [61, 62]. Comparing this ligand to other bibenzyl compounds in the library establishes the importance of a hydroxyl group located in the ethylene linker (Table 5). This information is used to guide construction of secondary libraries screened to improve the affinity and specificity of binding to PAS targets.

TABLE 5 Structure-activity information on PAS-A binding compounds. A.

B. compound R1 R2 K_(d) #134

—H ~100–200 μM #130

—CH₃ (>5 mM) #133

—H (>5 mM)

Comprehensive biochemical HTS for identifying specific, small molecule activators and inhibitors of PASK. We have adapted PASK protein binding assays, such as kinase and NMR-based assays, to high throughput screening (HTS). A compound collection initially consisting of roughly 350,000 drug-like chemicals has been collected, organized and extensively characterized in over 100 independent HTS assays. Such efforts have enabled the discovery of a large number of chemicals that potently and selectively modulate the activities of a broad range of polypeptide targets. The compound library has produced numerous drug entities that, following extensive optimization by medicinal chemistry, are in various phases of clinical and pre-clinical testing.

PASK HTS kinase assays identify both agonists and antagonists of the catalytic ability of the enzyme to phosphorylate the AMARA peptide. Our evidence indicates that the PASK PAS domains function as molecular sensors, negatively regulating the enzyme under conditions that are susceptible to depression in response to the appropriate intracellular signal. This parallels the AMPK system, which uses a regulatory domain and two negatively acting subunits to repress catalytic activity and simultaneously serve as the sensing device that monitors intracellular AMP/ADP-ATP ratios [4]. PAS domains have been reported to act as biosensors in studies of the aryl hydrocarbon receptor [73], photoactive yellow protein of the phototrophic bacterium E. halophila [74] and the bacterial Aer [75] and FixL [76] proteins.

One HTS assay kinase assay we employ is a robust and simple, non-radioactive chemiluminescent assay that measures the ability of the human enzyme to phosphorylate a biotinylated derivative of the AMARA peptide. The enzyme is expressed in baculovirus and purified by affinity and ion-exchange chromatography under routine conditions. Assay buffer is composed of 20 mM Tris-HC1, pH 7.5, 10 mM MgC1₂, 2 mM EGTA, 0.1% NP-40, 0.5 mM benzamidine, 1 mM NaF and 10 mM DTT. Enzyme is added to this buffer at a concentration determined to be in the linear range of activity (10 ng per microtiter plate well). For the most robust and sensitive kinase assay, we use an ATP concentration matching the K_(m)for PASK (2.5 mM). After dispensing 80 ul of the enzyme/buffer mix per well in Costar 96-well microtiter plates (coated with Neutravidin at 0.5 mg/ml at room temperature for 2 hr), 10 μl of test compounds is added at a final concentration of 1 μM compound. Compounds are diluted in DMSO from master plates stored in DMSO at 10 mM. Following compound addition, the reactions are initiated by addition of 10 μl of biotinylated AMARA peptide and ATP such that the final concentration of substrate and ATP are 1 mM and 2.5 mM respectively. Instrumentation for the addition of buffer/enzyme mix, diluted compounds and substrate/ATP is the Robbins Hydra 96 multipipette. Following brief shaking on a microtiter plate shaker (Lab-Line Instruments), plates are incubated for 1 hr at room temperature, washed 3× with distilled H₂O using a LABTEK 96-well wash, and applied with 100 μl per well of antibody buffer. The primary antibody is a mouse monoclonal antibody specific to the phosphorylated AMARA peptide (Pan Vera). The secondary antibody is an HRP-labeled rabbit, anti-mouse antibody (New England Biolabs). The final antibody buffer mix contains 2% BSA in PBS. After antibody addition the plates are briefly shaken, incubated at room temperature for 1 hr and washed 3× with H₂O. Reactions are developed by addition of 100 μl of Super Signal substrate (Pierce) and read colorimetrically using a Wallac Victor2, 1420 Multilabel Counter. Target specificity is confirmed by comparative assays with control non-PASK protein kinases.

Compounds showing good dose response curves, corresponding to potent agonistic or antagonistic activity, are tested against six versions of the human PASK enzyme. Three of the enzyme samples represent the intact protein in: (i) its native state as purified from baculovirus; (ii) its activated state following autophosphorylation; (iii) and its inactivated state following treatment with lambda phosphatase. Concomitant experiments are conducted using the amino-terminal truncated enzyme lacking its two PAS domains, again using the native enzyme as well as the phosphorylated and dephosphorylated derivatives. This secondary level of follow-up distinguishes between compounds that act on the kinase domain versus those that require the PAS domains for either stimulatory or inhibitory activity. Compounds from the HTS screen are evaluated by HPLC, mass spectrometry and NMR spectroscopy, not only of the starting compound, but also of analogs produced as a means of establishing a structure activity relationship (SAR) between the compound and its ability to either inhibit or stimulate PASK.

High throughput NMR-based protein/ligand screening. Two important facets of NMR-based protein/ligand screens are the size and scope of the compound libraries that are being used. One type of library we use is a “directed” library, consisting of a small (100–200) number of compounds whose structures are based on common structural elements of PAS-binding ligands, including biological cofactors (heme, NADH, FMN, FADH₂), chromophores (hydroxycinnaminic acid), environmental toxins (tetrachlorodibenzodioxin [TCDD]), etc. These have several features in common, including planarity, conjugation and limited charge at neutral pH. Chemical databases are searched with these criteria, starting with substructures that match common frameworks of PAS ligands. Additional geometric criteria are obtained from observations that protein-binding ligands are inherently biased towards certain chemical architectures [61, 79]. Many of these architectures are aromatic-rich and include the general structures of several PAS-binding ligands including PASK-binding compound #134 (Table 5). We also screen larger libraries (1000–2000 compounds) that are designed to cover a wider range of chemical structures while still taking advantage of the observed biases of protein-binding ligands [61, 79]. Compounds in this library are also chosen with an emphasis towards later use in synthetic approaches, with relatively low formula weight (100–200 Da) and composition of functional groups.

A wide variety of NMR-based methods are available to rapidly screen libraries of small compounds for binding to protein targets [82]. We primarily use protein-based screening methods, which are well suited for PAS domains given their small size (10–15 kDa) and the structural information obtained from the studies outlined above. As demonstrated with PAS-A, we screen using isotopically labeled protein and unlabeled ligands, using HSQC-type experiments to selectively observe protein signals in the presence of excess ligand. Compounds are maintained as 1M stocks in deuterated DMSO, and protein samples checked to ensure that DMSO does not bind with any significant affinity. Samples of ¹⁵N/¹³C-methyl labeled protein (250 μM) are mixed with 3–10 compounds at 1 mM each, which is sufficient to find ligands with weak (millimolar) dissociation constants. Protein chemical shifts are recorded using ¹³C/¹H HSQC as our primary method, complemented by ¹⁵N/¹H HSQC. Spectra from ligand-containing solutions are compared to those from ligand-free samples, calculating chemical shift changes with the minimum chemical shift difference method [83]. Where significant changes are observed, we deconvolute the binding of each ligand in the mixture by examining new samples with single protein/ligand mixtures. Compounds that demonstrate binding are titrated into a sample of ¹⁵N-labeled protein to measure dissociation constants, which can be measured by NMR if the complex is in fast exchange (K^(d)>10 μM).

Screening throughput of this method can exceed 1000 compounds per day for the initial screen: 15 minutes per sample (10′ experimental, 5′ sample manipulation) and 10 ligands/sample. PAS-binding compounds identified in this screen are evaluated for their ability to modulate PASK function in vitro using the previously described peptide phosphorylation assay. Compounds are checked for effects on both full length and truncated versions of PASK containing only the kinase domain to ensure that any modulatory action is mediated through the PAS domains.

We use several sources of structure-activity relationship information available from the first round of screening to generate small secondary libraries that we screen to find compounds with higher affinity. As shown above, comparisons of the affinity of structurally-related compounds identify positions on ligands that are essential for binding. Additionally, comparisons of the chemical shift changes caused by related ligands rapidly identify the relative orientation of protein/ligand complexes [85], potentially identifying sites on these ligands that may be amenable to modification as an avenue to increase affinity. The combination of these approaches allows us to design libraries that are simple modifications of ligands from the first screen.

An alternative approach that generates more drastically altered ligands for the second library is the SAR-by-NMR method [86]. We conduct two screens through the primary library with the goal of finding two ligands that bind at adjacent sites on the protein structure. This is achieved by a first screen as previously discussed, followed by a second screen through the library in the presence of saturating concentrations of a ligand identified by the first screen. Through the use of protein-directed NMR experiments such as the ¹³C/¹H HSQC, one can readily identify compounds from the second screen that bind at an adjacent site to the first compound. We then obtain a structure of the ternary complex of the protein bound to both ligands. This is achieved by obtaining similar structural information as used for the PAS-A structure determination complemented by intermolecular distance restraints using isotope-filtered, edited 3D NOE experiments [87] that provide the selective observation of NOEs between isotopically-enriched proteins and non-enriched ligands. Guided by this structure, we identify sites on each ligand that can be used to generate a linker between the two compounds in the expectation that the linked compound will have enhanced affinity over the two separate compounds. Libraries are constructed from this information using a series of linkers with variable length and flexibility. We screen these secondary library compounds for binding to the PAS-A and PAS-B domains as described above.

Phosphorylation Screening. An alternative strategy to identify PASK substrates follows methods developed by Hunter and colleagues for the identification of protein kinase substrates by expression screening with solid-phase phosphorylation. This method, termed “phosphorylation screening,” represents an adaptation of classical methods of expression screening using bacteriophage lambda. Hunter and colleagues have used this approach to identify known and novel substrates for the Erk1 [65] and Cdk2 [66] protein kinases.

Yeast cDNA are cloned into the λGEX5 vector developed by Hunter and colleagues. This bacteriophage lambda vector contains a plasmid sequence between two NotI sites, consisting of a Co1E1 origin, the ampicillin resistance gene, and a GST gene followed by a small “stuffer” sequence to be replaced by cDNA inserts. Compared with the original λgt11 vector, the λGEX5 vector provides three important advantages. First, clones encoding polypeptides phosphorylated by PASK can be rapidly converted into plasmid clones by excision rescue without purifying cDNA fragments. Second, rescued plasmids can be directly utilized not only for cDNA sequencing but also for expression of GST fusion proteins for further characterization. Finally, GST, the N-terminal fusion partner of the recombinant product expressed by λGEX5, is highly soluble, easy to purify by GSH-agarose chromatography, and much smaller than the β-galactosidase component of the fusion proteins expressed by λgt11.

Expression libraries are screened according to the methods of Hunter and Fukunaga [65] using human PASK expressed in baculovirus. Prior to the phosphorylation step of the assay, filter lifts are incubated in the presence of unlabelled ATP as a means of reducing the frequency of isolating clones whose products have either autophosphorylating or ATP-binding activity. Highly purified, auto-phosphorylated (activated) human PASK is added with [γ-³²P]ATP in optimal buffer conditions. Following incubation, the filters are washed and exposed to X-ray film to identify plaques expressing polypeptide substrates for PASK. Optimization strategies outlined by Hunter and Fukunaga are followed to expedite completion of a successful screen. After retrieving nitrocelluose lifts bearing lambda-expressed proteins, filters are exposed to 6M urea followed by sequential dilution and ultimate removal of the urea. This approach helps solubilize over-expressed proteins which are often restricted to inclusion bodies. In the case of expression cloning of eukaryotic transcription factors, inclusion of the urea solubilization step substantially enhanced signal to noise ratios [67].

Positive plaques are isolated, from which encoding plasmids are excised and sequenced. Particularly interesting substrate proteins are further studied to identify precise sites of phosphorylation using GST-fusion proteins expressed and purified from E. coli. Finally, individual serine and threonine sites are altered by site directed mutagenesis followed by reintroduction of the mutant on a plasmid in a strain deleted for the chromosomal copy of the gene so as to facilitate further biochemical and biological assays.

In addition to phosphorylation screening, we employ an entirely different approach described by Kirschner and colleagues [68–70]. This approach also relies on the screening of cDNA libraries, yet in this case the assay is conducted on pools of in vitro translation products. In brief, a cDNA library is prepared from poly(A)+ yeast RNA cloned into the pCS2+ vector [71]. cDNAs larger than 0.5 kb is ligated directionally into pCS2+ and electroporated into competent E. coli cells. The pCS2+ plasmid contains regulatory sequences for coupled transcription/translation immediately upstream from the directionally cloned cDNA insert site, thus facilitating TNT-mediated transcription/translation in reticulocyte lysates. We follow the methods of Kirshner and colleagues to simultaneously express pools of ˜100 cDNA clones, typically resulting in the synthesis of 15–30 ³⁵S-labeled polypeptides. These pools of in vitro translated protein will subsequently be incubated with baculovirus-expressed PASK under optimal conditions for its enzymatic activity. Following phosphorylation with cold ATP, the ³⁵S-labeled protein pools are run on SDS-PAGE gels followed by autoradiographic imaging. Control reactions are conducted in the absence of PASK such that pooled samples can be compared side-by-side on SDS-PAGE gels. Proteins phosphorylated by PASK are provisionally identified by a shift in their electrophoretic mobility. Pools containing cDNA capable of expressing PAS-kinase modified polypeptides are de-convoluted in order to identify relevant cDNAs and their encoded polypeptides.

Protocol for High Throughput PASK Autophosphorylation Assay.

A. Reagents:

-   -   Neutralite Avidin: 20 μg/ml in PBS.     -   kinase: 10⁻⁸–10⁻⁵M PASK kinase domain at 20 μg/ml in PBS.     -   substrate: 10⁻⁷–10⁻⁴M biotinylated PASK substrate at 40 μg/ml in         PBS.     -   Blocking buffer: 5% BSA, 0.5% Tween 20 in PBS; 1 hour at room         temperature.     -   Assay Buffer: 100 mM KCl, 10 mM MgCl₂, 1 mM MnCl₂, 20 mM HEPES         pH 7.4, 0.25 mM EDTA, 1% glycerol, 0.5% NP-40, 50 mM BME, 1         mg/ml BSA, cocktail of protease inhibitors.     -   [³²P]γ-ATP 10× stock: 2×10⁻⁵M cold ATP with 100 μCi [³²P]γ-ATP.         Place in the 4° C. microfridge during screening.     -   Protease inhibitor cocktail (1000×): 10 mg Trypsin Inhibitor         (BMB # 109894), 10 mg Aprotinin (BMB # 236624), 25 mg         Benzamidine (Sigma # B-6506), 25 mg Leupeptin (BMB # 1017128),         10 mg APMSF (BMB # 917575), and 2 mM NaVo₃ (Sigma # S-6508) in         10 ml of PBS.         B. Preparation of Assay Plates:     -   Coat with 120 μl of stock N Avidin per well overnight at 4° C.     -   Wash 2 times with 200 μl PBS.     -   Block with 150 μl of blocking buffer.     -   Wash 2 times with 200 μl PBS.         C. Assay:     -   Add 40 μl assay buffer/well.     -   Add 40 μl biotinylated substrate (2–200 pmoles/40 ul in assay         buffer)     -   Add 40 μl kinase (0.1–10 pmoles/40 ul in assay buffer)     -   Add 10 μl compound or extract.     -   Add 10 μl [³²P]γ-ATP 10× stock.     -   Shake at 25° C. for 15 minutes.     -   Incubate additional 45 minutes at 25° C.     -   Stop the reaction by washing 4 times with 200 μl PBS.     -   Add 150 μl scintillation cocktail.     -   Count in Topcount.         D. Controls for All Assays (Located on Each Plate):     -   a. Non-specific binding     -   b. cold ATP at 80% inhibition.

Protocol for High Throughput PASK PAS Domain-Ligand Binding Assay.

A. Reagents:

-   -   Neutralite Avidin: 20 μg/ml in PBS.     -   Blocking buffer: 5% BSA, 0.5% Tween 20 in PBS; 1 hour at room         temperature.     -   Assay Buffer: 100 mM KCl, 20 mM HEPES pH 7.6, 1 mM MgCl₂, 1%         glycerol, 0.5% NP-40, 50 mM b-mercaptoethanol, 1 mg/ml BSA,         cocktail of protease inhibitors.     -   ³³P PASK polypeptide 10× stock: 10⁻⁸–10⁻⁶ M “cold” PASK PAS         domain supplemented with 200,000–250,000 cpm of labeled PASK         (Beckman counter). Place in the 4° C. microfridge during         screening.     -   Protease inhibitor cocktail (1000×): 10 mg Trypsin Inhibitor         (BMB # 109894), 10 mg Aprotinin (BMB # 236624), 25 mg         Benzamidine (Sigma # B-6506), 25 mg Leupeptin (BMB # 1017128),         10 mg APMSF (BMB # 917575), and 2 mM NaVO₃ (Sigma # S-6508) in         10 ml of PBS.     -   Ligand: 10⁻⁷–10⁻⁵M biotinylated ligand in PBS.         B. Preparation of Assay Plates:     -   Coat with 120 μl of stock N-Avidin per well overnight at 4° C.     -   Wash 2 times with 200 μl PBS.     -   Block with 150 μl of blocking buffer.     -   Wash 2 times with 200 μl PBS.         C. Assay:     -   Add 40 μl assay buffer/well.     -   Add 10 μl compound or extract.     -   Add 10 μl ³³P-PASK polypeptide (20–25,000 cpm/0.1–10         pmoles/well=10⁻⁹–10⁻⁷ M final conc).     -   Shake at 25° C. for 15 minutes.     -   Incubate additional 45 minutes at 25° C.     -   Add 40 μM biotinylated ligand (0.1–10 pmoles/40 ul in assay         buffer)     -   Incubate 1 hour at room temperature.     -   Stop the reaction by washing 4 times with 200 μM PBS.     -   Add 150 μM scintillation cocktail.     -   Count in Topcount.         D. Controls for All Assays (Located on Each Plate):     -   a. Non-specific binding     -   b. Soluble (non-biotinylated ligand) at 80% inhibition.

Literature Cited.

-   1. Hanks, S. K., et al. Science, 1988. 241(4861):42–52. -   2. Dale, S., et al. FEBS Letters, 1995. 361(2–3):191–5. -   3. Stein, S. C., et al. Biochemical J, 2000. 345(Pt 3):437–43. -   4. Hardie, D. G., et al. Annual Review of Biochemistry, 1998.     67:821–55. -   5. Crute, B. E., et al. J Biological Chemistry, 1998.     273(52):35347–35354. -   6. Woods, A., et al. Molecular & Cellular Biology, 2000.     20(18):6704–6711. -   7. Hampsey, M., Yeast, 1997. 13(12):1099–133. -   8. Sikorski, R. S., et al. Genetics, 1989. 122(1 ):19–27. -   9. Scriver, S. C., et al. The Metabolic Basis of Inherited Disease     (6th ed). 1989:ch. 13. -   10. Boles, E., et al. European J Biochemistry, 1994. 220(1):83–96. -   11. Mylin, L. M., et al. Genetics, 1994. 137(3):689–700. -   12. Cartwright, J. L., et al. J Biological Chemistry, 1999.     274(13):8604–10. -   13. Safrany, S. T., et al. J Biological Chemistry, 1999.     274(31):21735–40. -   14. Chuang, R. Y., et al. Science, 1997. 275(5305):1468–71. -   15. Jamieson, D. J., et al. Molecular Microbiology, 1991.     5(4):805–12. -   16. Jamieson, D. J., et al. Nature, 1991. 349(6311):715–7. -   17. Dunckley, T., et al. EMBO Journal, 1999. 18(19):5411–22. -   18. Chu, S., et al. Rna, 1997. 3(4):382–91. -   19. Lee, J. Y., et al. Molecular & Cellular Biology, 1991.     11(2):721–30. -   20. Mendenhall, M. D., et al. Nucleic Acids Research, 1988.     16(17):8713. -   21. Venema, J., et al. Annual Review of Genetics, 1999. 33:261–311. -   22. Grishin, N. V., Trends in Biochemical Sciences, 1998.     23(9):29–30. -   23. Lim, A. L., et al. J Biological Chemistry, 1996.     271(19):11400–11409. -   24. Werner, M., et al. Cell, 1987. 49(6):805–13. -   25. Wang, Z., et al. J Biological Chemistry, 1999. 274(53):37565–74. -   26. Volarevic, S., et al. Science, 2000. 288(5473):2045–7. -   27. Lazaris-Karatzas, A., Nature, 1990. 345(6275):544–7. -   28. Clemens, M. J., et al. International J Biochem & Cell     Biol, 1999. 31(1):1–23. -   29. De Benedetti, A., et al. International J Biochem & Cell     Biol, 1999. 31(1):59–72. -   30. Nathan, C. A., et al. Oncogene, 1997. 15(9):1087–94. -   31. Nathan, C. A., et al. Oncogene, 1997. 15(5):579–84. -   32. Li, B. D., et al. Cancer, 1997. 79(12):2385–90. -   33. Miyagi, Y., et al. Cancer Letters, 1995. 91(2):247–52. -   34. Anthony, B., et al. International J Cancer, 1996. 65(6):858–63. -   35. Eberle, J., et al. International J Cancer, 1997. 71(3):396–401. -   36. Brass, N., et al. Human Molecular Genetics, 1997. 6(1):33–9. -   37. Asano, K., et al. JBiological Chemistry, 1997. 272(38):23477–80. -   38. Raught, B., et al. International J Biochem & Cell Biology, 1999.     31(1):43–57. -   39. Liang, L., et al. Development, 1994. 120(5):1201–11. -   40. Cuff, J. A., et al. Bioinformatics, 1998. 14:892–893. -   41. Sheffield, P., Protein Expression and Purification, 1999.     15:34–39. -   42. Gronenborn, A. M., et al Protein Science, 1996. 5:174–177. -   43. Huth, J. R., et al. Protein Science, 1997. 6:2359–2364. -   44. Clore, G. M., et al. Trends Biotechnol., 1998. 16:22–34. -   45. Sattler, M., et al. Progr Nucl Magnetic Resonance     Spectrosc, 1999. 34:93–158. -   46. Neri, D., et al. Biochemistry, 1989. 28:7510–7516. -   47. Pascal, S. M., et al. J Magnetic Resonance B, 1994. 103:197–201. -   48. Fletcher, C. M., et al. J Biomolecular NMR, 1996. 8:292–310. -   49. Cornilescu, G., et al. J Biomolecular NMR, 1999. 13:289–302. -   50. Brünger, A. T., et al. Acta. Cryst., 1998. D54:905–921. -   51. Nilges, M., et al. Progr Nucl Magnetic Resonance Spectrosc,     1998.32:107–139 -   52. Hansen, M. R., et al. Nature Structural Biology, 1998.     5(12):1065–1074. -   53. Borgstahl, G. E. O., et al. Biochemistry, 1995. 34:6278–6287. -   54. Cabral, J. H. M., et al. Cell, 1998. 95:649–655. -   55. Düx, P., et al. Biochemistry, 1998. 37(37):12689–12699. -   56. Gong, W., et al. Proc. Natl. Acad. Sci., 1998. 95:15177–15182. -   57. Farrow, N. A., et al. Biochemistry, 1994. 33:5984–6003. -   58. Bracken, C., et al. J Molecular Biology, 1999. 285:2133–2146. -   59. Farrow, N. A., et al. J Biomolecular NMR, 1995. 6:153–162. -   60. Pellequer, J. L., et al. Current Biology, 1999. 9(11):R416–8. -   61. Fejzo, J., et al. Chemistry and Biology, 1999. 6:755–769. -   62. Hajduk, P. J., et al. J Medicinal Chemistry, 2000. 43(21):3862–6 -   63. Kranz, J. E., et al.Proc. of the Nat.l Acad. Sci. USA, 1990.     87(17):6629–33. -   64. Koshland, D., et al. Cell, 1985. 40(2):393–403. -   65. Fukunaga, R., et al. EMBO Journal, 1997. 16(8):1921–33. -   66. Jiang, W., et al. Molecular Cell, 1998. 2(6):877–85. -   67. Vinson, C. R., et al. Genes & Development, 1988. 2(7):801–6. -   68. Stukenberg, P. T., et al. Current Biology, 1997. 7(5):338–48. -   69. Lustig, K. D., et al. Methods in Enzymology, 1997. 283:83–99. -   70. King, R. W., et al. Science, 1997. 277(5328):973–4. -   71. Turner, D. L., et al. Genes & Development, 1994. 8(12):1434–47. -   72. McKnight, S. L., et al. Science, 1982. 217(4557):316–24. -   73. Rowlands, J. C., et al. Critical Reviews in Toxicology, 1997.     27(2):109–34. -   74. Farber, G. K., Nature Structural Biology, 1998. 5(6):415–7. -   75. Taylor, B. L., et al. Annual Review of Microbiology, 1999.     53:103–28. -   76. Pellequer, J. L., et al. CurrBiol, 1999. 9(11):R416–8. -   77. Wang, Z., et al. Structure, 1998. 6(9):1117–28. -   78. Gum, R. J., et al. J Biological Chemistry, 1998.     273(25):15605–10. -   79. Hajduk, P. J., et al. J American Chemical Society, 2000.     122:7898–7904. -   80. Matthews, S. J., et al. J Biomolecular NMR, 1993. 3:597–600. -   81. Meiler, J., et al. J Biomolecular NMR, 2000. 16:245–252. -   82. Hajduk, P. J., et al. Quarterly Reviews of Biophysics, 1999. 32     (3): 211–40 -   83. Farmer, B. T. I., et al. Nature Structural Biology, 1996.     3:995–997. -   84. Johnson, B. A., et al. J Biomolecular NMR, 1994. 4:603–614. -   85. Medek, A., et al. J American Chemical Society, 2000.     122(6):1241–1242. -   86. Shuker, S. B., et al. Science, 1996. 274:1531–1534. -   87. Zwahlen, C., et al. J American Chemical Society, 1997.     119:6711–6721.

All publications and patent applications cited in this specification are herein incorporated by reference as if each individual publication or patent application were specifically and individually indicated to be incorporated by reference. Although the foregoing invention has been described in some detail by way of illustration and example for purposes of clarity of understanding, it will be readily apparent to those of ordinary skill in the art in light of the teachings of this invention that certain changes and modifications may be made thereto without departing from the spirit or scope of the appended claims. 

1. A composition comprising an isolated polynucleotide encoding a polypeptide wherein the polypeptide comprises SEQ ID NO:2.
 2. A composition according to claim 1, wherein the polynucleotide comprises SEQ m NO:
 1. 3. An expression vector comprising a polynucleotide according to claim
 1. 4. A cell comprising a polynucleotide according to claim 1, wherein the polypeptide is expressed in the cell.
 5. A composition comprising an isolated polynucleotide encoding a polypeptide wherein the polypeptide comprises residues 1–89 of SEQ ID NO:2 and is sufficient to elicit a human PASK (SEQ ID NO:2)-specific antibody in a heterologous host.
 6. A composition according to claim 5, wherein the polynucleotide comprises nucleotides 391–711 of SEQ ID NO:1.
 7. A composition according to claim 5, wherein the polynucleotide comprises nu.cleotides 1021–1317 of SEQ ID NO:
 1. 8. An expression vector comprising a polynucleotide according to claim
 5. 9. A cell comprising a polynucleotide according to claim 5, wherein the polypeptide is expressed in the cell. 